Technovit H7100/H8100

Staining Protocol for Lymphoid Tissue

Normal immunostaining procedures for routine markers on sections obtained from lymphoid tissue.

Procedure

1. Dry the sections for two hours at 37°C on a slide warmer. This must be done whether the slides have been stored at 4°C for some time or just collected.
2. Pretreat the sections with trypsin at 37°C. Trypsin concentrations must be determined separately for each antibody. Use trypsin solution that has been preheated at 37°C for 30 minutes. Cover section with at least 100 micro litres of solution.
3. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times.
4. Preincubate in normal serum from the animal species in which the second antibody is raised for 30 minutes at 37°C. Only perform this step if aspecific background is present using a particular antibody.
5. Drip off excess serum and apply the first antibody in an appropriate concentration and incubate for two hours at 37°C.
6. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times.
7. Block endogenous preoxidase in a solution of 0.05% hydrogen peroxide in phosphate buffered saline, pH 7.4, for 30 minutes at room temperature.
8. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times.
9. Incubate in appropriate dilutions of the secondary antibody, containing 5% normal serum for 60 minutes at room temperature.
10. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times.
11. Develop the peroxidase activity in daminobenzidine (DAB).
12. Counterstain the sections in Haematoxylin or if a more advanced morphological detail is necessary, in periodic‐acid‐Schiff reagent.
13. Cover with Glycerin‐gelatin and cover glass.

Sectioning and Mounting Protocol

Sectioning

Sectioning is best done with a rotary or sledge microtome such as the JB‐4 Microtome. The features to look for are retraction of the specimen on the return stroke and motorised motion of the sample relative to the knife edge. Retraction is needed to keep the hard sample from brushing against the knife edge on the return stroke, damaging the edge and block face. Motorised sample movement is desirable to get more reproducible cutting speed and force.

The best sections are obtained on these microtome with a glass knife. Glass knife strips are available from Electron Microscopy Sciences. Dry block faces are used; in fact, moisture and humidity will soften GMA blocks sufficiently to make sectioning impossible. If the block is too soft to section dry the block in a warm oven or move to a less humid work environment.

Sections are collected from the knife edge using tweezers and stretched by floating on a water bath. Two factors are important when preparing the water bath: cleanliness and temperature. Any residue of soap or oil will decrease the surface tension of the water dramatically reducing its stretching ability and/or causing the section to sink. Increasing temperature also reduces the stretching ability of the bath so that at 20°C stretching of 10‐13% is possible while at 60°C stretching of 7‐9% is possible. Section stretching allows recovery of almost all the compression caused during sectioning. Properly stretched, much less than 1% of the compression in vertical dimension remains in the sections.

Most procedures call for mounting your sections on slides before staining. The exception may be when staining sawn sections since these retain enough strength to be held in tweezers and hand‐dipped in the various staining solution (followed directly by coverslipping to slides). Mounting thinner sections to slides before staining also helps to prevent folds from developing in the sections. If you are using slides precoated with silane, skip the protocol for coating your own.

Preparing Silanised Slides

Procedure

1. Dip slides in 2% silane/acetone solution for one minute.
2. Dip slides in 100% acetone for one minute.
3. Dip slides in double distilled water for one minute.
4. Repeat step 3 with agitation.
5. Air dry.
6. Apply sections on a drop of water on the slide.
7. Dry on 60°C hot plate for two to five minutes.
8. Air dry at room temperature overnight.

1. Dip your coated slide into the waterbath under the section and lift the section off the water surface. Dry the slides for about 16 minutes on a 60°C slide warmer.

Prussian Blue Stain for Glycol Methacrylate Sections

Procedure

1. Stain in Potassium Ferricyanide solution at 60°C for 15 minutes. Filter solution after heating and before use.
2. Wash in distilled water.
3. Stain in Safranin O for two to five minutes.
4. Wash in 1% acetic acid.
5. Dehydrate in 96% alcohol two times then 100% alcohol.
6. Clear in xylene and mount.

Results

Solutions

Potassium Ferricyanide Solution:

• Potassium Ferricyanide ‐ 1g
• Distilled water ‐ 50ml
• 2% Hydrochloric acid ‐ 50ml

Safranin O:

• Safranin O (C.I. #50240) ‐ 0.2g
• 1% acetic Acid ‐ 100ml

Citation

Gerrits, P.O. and Smid, L., "Staining Procedures for Tissues Embedded in 2‐Hydroxyethyl Methacrylate", Heraeus Kulzer

Periodic Acid Schiff Stain for Glycol Methacrylate SectionsProcedure

1. Oxidise sections in 0.4% periodic acid for 30 minutes at 57°C.
2. Wash in running tap water or sodium sulphite solution (to avoid a general pink background).
3. Rinse in three times in distilled water.
4. Schiff's Reagent for 15 minutes.
5. Wash well in running tap water.
6. Rinse in distilled water.
7. Counterstain with Gill's Haematoxylin for 10 minutes.
8. Wash well in running tap water.
9. Dehydrate using 95% and 100% ethanol.
10. Clear in xylene and cover slip.

Results

Solutions

Gill's Haematoxylin:

• Haematoxylin (C.I. 75290) ‐ 6g
• Sodium Iodate ‐ 0.6gm
• Aluminium Sulphate ‐ 52.8g
• Distilled water ‐ 690ml
• Ethylene Glycol ‐ 250ml
• Glacial acetic acid ‐ 60ml

Schiff's Reagent:

• Mix 0.5g of Pararosaniline (C.I.#42500) into 15mL of 1N Hydrochloric Acid.
• Mix 0.5g of Potassium Metabisulphite into 85ml of distilled water.
• Add the second solution to the first and store for 24 hours in a dark place.
• Bleach the resulting solution with 200mg of bone charcoal and filter.
• Store the colourless solution at 4°C.

Sodium Sulphite Solution:

• 0.5g of sodium sulphite
• 100ml of distilled water

Haematoxylin and Eosin Stain for Glycol Methacrylate Sections

Haematoxylin and Eosin stain is a good general stain for many types of tissue.

Solutions

Gills Haematoxylin

• 6g Haematoxylin (C.I. 75290)
• 0.6g Sodium Iodate
• 52.8g Aluminium Sulphate
• 690ml Distilled Water
• 250ml Ethylene glycol
• 60ml Glacial acetic Acid

Eosin

• 0.5g Eosin Y (Alcoholic C.I. 45380)
• 100ml Ethanol 96%
• 2 drops Glacial Acetic Acid

Acid Ethanol

• 4ml 0f 25% HCl
• 100ml of 70% Ethanol

Procedure

1. Stain in Gill's Haematoxylin for 15 minutes.
2. Wash in tap water for 10 minutes.
3. Decolorize plastic, if necessary, with acid alcohol followed by tap water rinse.
4. Rinse in distilled water.
5. Scott's Tap Water for one to two minutes.
6. Two changes of distilled water.
7. Counterstain with Eosin for two to five minutes.
8. Dehydrate using 96% and 100% ethanol.
9. Clear in xylene and cover slip.

Results

Glycol Methacrylate Embedding for Materials Samples

Embedding in plastic often supplies the support necessary to successfully section various materials samples, especially prousand inhomogeous samples. GMA has been used to embed and section polymer resin, multilayer foils and films, paper, textiles and coatings in addition to a wide variety of biological material. The ability of this low viscosity resin to infiltrate effectively can make sectioning non‐biological samples fast and economical.

Preparation of this type of sample is slightly different than preparing biological samples.

Procedure

A fixation step is, of course, not necessary for materials samples.

Dehydration for GMA embedments does not have to be complete because of GMA's miscibility in water. Only if the sample is saturated with water should a schedule of increasing alcohol concentration be used. This will displace most of the water with a solvent that can be replaced with plastic. Use schedule as follows:

• 70% ethanol for 10minutes
• 96% ethanol for 10 minutes
• 96% ethanol for 10 minutes
• Absolute ethanol for 10 minutes

Infiltration displaces the dehydration solvent with monomer prior to beginning the polymerisation reaction. If you have a porous sample, infiltration may be needed. Nonporous samples can skip this step. Make up your infiltrating solution according to the directions from your kit. While complete elimination of air is not needed, sealing the moulds with paraffin or plastic wrap is often done. Leave the samples at room temperature for one hour then at 37°C for one hour to complete polymerisation.

If your embedments cannot be mounted directly to the microtome, an adapter must be attached. Place the suitable Technovit 3040 is an inexpensive media prepared by mixing three parts of the dry component to one part of the liquid component. Pour it into the adapter and polymerization will be complete with 10 minutes. The adapters will be firmly attached through a copolymerization of the Technovit 3040 with the GMA.

You are now ready to begin sectioning with glass.

Giemsa Stain For Glycol Methacrylate Sections

This Giemsa stain works well for a variety of tissues.

Procedure

1. Stain in Giemsa working solution at room temperature for 1½ hours.
2. Rinse in dilute acetic acid (four drops in 100mL DI water) for two seconds.
3. Dip in 96% alcohol.
4. Dip in 96% alcohol.
5. Rinse in isopropyl alcohol three times for two minutes each.
6. Clear in xylene and mount.

Results

Solutions

Giemsa Solution:

• 0.15g Azure A
• 0.30g Methylene Blue
• 0.36g Eosin Y
• 0.04g Phloxine B
• 50ml Glycerine
• 40ml Methanol
• Add dyes to Glycerine and methanol, mix well. Incubate overnight in 55‐58°C oven stirring occasionally. Stable for year.

Giemsa Working Solution:

• 4mL Giemsa stock solution
• 40mL phosphate buffer solution

Phosphate Buffer Solution:

• 7.25g Sodium phosphate, monobasic
• 2.75g Sodium phosphate, dibasic
• 1000ml Distilled water
• 10ml 10% Triton X‐100
• Store at 4°C.

Glycol Methacrylate Embedding for Soft Tissues

Embedding in plastic provides many advantages to the histotechnologist. Thinner sections can be made providing improved detail. Better support is given to cellular components, offering improved morphology. Short, straightforward protocols are available giving minimum processing times. Improved chemistry provided by kits gives uniform results and a wider range of application.

While this protocol will help get you started with embedding your soft biological samples in the Technovit GMA kits sold by Electron Microscopy Sciences, it follows the general guidelines for embedding in any source of GMA.

Procedure

Fixation of the biological samples can be done in any way appropriate for the work you are doing. Immersion or perfusion with 4% neutralised formaldehyde, prepared from paraformaldehyde according to Karnovsky, is usually preferred. Try to keep the sample size small using 10mm x10mm 2xmm as a maximum.

Dehydration for GMA embedments does not have to be complete because of GMA's miscibility in water. Use a schedule of increasing alcohol concentration at room temperature as follows:

• 70% ethanol for two hours
• 96% ethanol for two hours
• 96% ethanol for two hours
• Absolute ethanol for one hour

A short defatting step can help with infiltration. If desired, submerge the sample in acetone for 10 minutes.

Make your infiltrating solution from:

• 100ml of Technovit 7100 resin, i.e. 2‐hydroxyethyl methacrylate (GMA)
• 1g of Hardener I (benzoyl peroxide)

Mix using a magnetic stirrer until the benzoyl peroxide is complete dissolved. Store at 4°C in a dark bottle for up to two months.

Infiltrate in a 50/50 mixture of 100% ethanol and infiltration solution for two hours. Leave the sample in 100% infiltration solution overnight. Infiltration with mild agitation and/or vacuum will be more complete and larger samples should have more infiltration steps over a longer period of time.

Make up your embedding solution from:

• 15 parts of infiltrating solution
• 1 part of Hardener II

Mix for one minute using a magnetic stirrer. Use the solution within 10 minutes, before polymerisation occurs.

Glycol Methacrylate Embedding For Immunohistochemistry

Embedding in plastic provides many advantages to the histotechnologist. Thinner sections can be made providing improved detail. Better support is given to cellular components, offering improved morphology. Short, straight forward protocols are available giving minimum processing times. In addition, improved chemistry and protocols are now available that are more antigen‐friendly.

This protocol will help get you started with embedding your soft biological samples in Technovit 8100, a GMA kit sold by Electron Microscopy Sciences. Note the areas in which it deviates from the guidelines for embedding in GMA for morphology only.

Procedure

1. Use gentle agitation during fixation, washing, dehydration and infiltration for best results.
2. Fixation of the biological samples can be done in any way appropriate for the antigen you are seeking. Cold acetone, periodate/lysine/paraformaldehyde, and buffered paraformaldehyde solutions have all been used successfully. Try to keep the sample size small using 10mmx10mm2mm as a maximum. Fixation should be done for several hours at 4°C.
3. Wash out the fixative with 6% sucrose solution at 4°C overnight.
4. Dehydration for GMA embedments does not have to be complete because of GMA's miscibility in water. Use acetone for one hour at 4°C changing the acetone at the beginning until it remains clear.
   Make up your infiltration solution from:
   100mL of Technovit 8100 resin, i.e. 2‐hydroxyethyl Methacrylate (GMA)
   0.6gm of Hardener I (benzoyl peroxide)
5. Mix using a magnetic stirrer until the benzoyl peroxide is complete dissolved. Store at 4°C in a dark bottle for up to one month.
6. Infiltrate for six to ten hours at 4°C using mild agitation. Vacuum will make infiltration more complete and larger samples should have more infiltration steps over a longer period of time.
   Make up your embedding solution from:
   30mL of infiltrating solution
   1mL of Hardener II
7. Mix for one minute using a magnetic stirrer. Use the solution within 10 minutes, before polymerisation occurs.
8. Cover your samples in the embedding moulds with this solution. Moulds of polyethylene such as the JB‐4 mould or Peel‐away moulds are most often used. Block holder/adapter in the recess on the mould and add mounting plastic.

Alkaline Phosphatase for Glycol Methacrylate Sections

Procedure

1. Incubate sections in the incubating medium at room temperature for one to three hours. Two hours is sufficient in most cases.
2. Wash in distilled water for two minutes.
3. Counterstain with Nuclear Fast Red for five to ten minutes.
4. Wash in distilled water for two minutes.
5. Air dry and coverslip.

Results

To preserve the reaction product, selection of the right mounting (coverslip) medium is important.

Solutions

Incubating Medium:

• 5mg Naphtol As‐MX phosphate, di‐sodium salt (sigma)
• 0.25ml N,N‐dimethylformamide
• 30mg Fast blue BB (sigma)
• 25ml Distilled water
• 25ml Buffer Solution
• 2 drops 10% Magnesium Sulphate Solution
  Prepare fresh, shake well and filter before use.

Buffer Solution:

• 2.4g 0.2M Tris (Hydroxyethyl)‐aminomethane
• 100ml Distilled water
• Adjust the pH of the buffer to 8.9 with dilute HCl and store at 4°C.

Nuclear Fast Red:

• To 0.2g of Nuclear Fast Red add 200ml of boiling 0.5% aluminium sulphate solution
• Keep boiling for five to ten minutes.
• Allow to cool and filter before use.

Acid Phosphatase for Glycol Methacrylate Section

Procedure

1. Incubate sections in the incubating medium at 37°C for five to 12 hours. Long incubation periods are needed to get significantly visible reaction product.
2. Wash in distilled water for two minutes.
3. Counterstain with Methyl Green for five minutes.
4. Wash in distilled water for two minutes.
5. Air dry and cover slip.

Results

Solutions

Incubating Medium:

• Combine 20ml of buffer solution, 48ml of distilled water and 4ml of substrate solution.
• Combine 3.2ml of Pararosaniline solution with 3.2ml of sodium nitrite solution. Mix for one minute.
• Add the second solution to the first.
• Adjust pH to 5.

Buffer Solution:

• 5.9 g Anhydrous sodium acetate
• 14.7g Sodium barbiturate
• 500ml Distilled water (boiled)

Do not adjust the pH of the buffer and store at 4°C.

Substrate solution:

• 40mg Naphtol As‐BI phosphatease, sodium salt
• 4ml N.N‐dimethylformamide

Pararosaniline Solution:

• 2g Pararosaniline (C.I.#42500)
• 50ml 2N HCl

Use heat to dissolve, filter when cool and store at 4°C

Sodium Nitrite Solution:

• Sodium Nitrite ‐ 1g
• Distilled Water ‐ 25ml

Prepare fresh and store at 4°C.

Methyl Green

• Methyl Green (C.I.# 42585) ‐ 1g
• Phosphate/citrate buffer 0.1M pH 4.0 ‐ 100ml

Citation:

Gerrits, P. O. and Smid, L., "Staining Procedures for Tissues Embedded in 2‐Hydroxyethyl Methacrylate", Heraeus Kulzer.